We specialize in developing new microscopy methods and harness the scientific insight that is often conceived at frontiers where diverse disciplines meet. In biology, several such fields are converging: new ways to label tissues at the molecular level, new techniques in optical and electron microscopy, powerful electronics to collect terabytes of data, and computer algorithms that can reveal complex patterns and relationships.

At JFRC, I lead a team that seeks to understand bioimaging challenges, to harness and introduce new disciplines or technologies that are relevant, and to promote a coherent systems approach for optimal benefit to biological research. Two initial projects will characterize this effort: higher-resolution 3D optical microscopy and high-throughput 3D electron microscopy.

Such ambitious projects can succeed only with the collective participation of researchers at JFRC. The availability of genetically engineered samples, customized labels, instrumentation support, computation infrastructure, image processing, and theoretical modeling—as well as biologists to give inspiration and direction—is critical. I hope to bridge diverse fields, nucleate and drive collaborations, and be driven by new research opportunities identified by scientists at JFRC.

The detail that can be seen by optical microscopy is limited to ~0.25 microns (about 0.25 percent the diameter of a human hair). Although this level of detail can resolve some of the organelles of a cell, it falls short of revealing all the details of an organelle and is even less able to show any nanometer-sized structure that is responsible for so much of molecular biology. Combining three insights has led us to a new kind of optical microscopy that can peer into this regime:

Single fluorescent molecules (which might label a protein of interest) can be imaged and their centers can be localized to a fraction of the size of the fuzzy spot that corresponds to their image.

Closely spaced, optically overlapping fluorescent molecules can be separated, and each can be localized if there is a distinguishing characteristic. For example, if two molecules light up separately in different image frames, the center of each can be inferred to a fraction of the spot sizes.

A new class of activatable fluorescent proteins has been developed in the past several years, and a distinguishing subset of these proteins can be turned into a fluorescing state while the remainder remain dark.

This last insight led to a new kind of microscope that my Janelia colleague and co-inventor, Eric Betzig, and I have dubbed PALM (photoactivated localization microscopy). Watch a video of the personal story of how we invented PALM. This microscope can activate, sample, and localize the centers of a very small subset of closely spaced label molecules. After bleaching the first subset, this process is repeated for a new sparse subset to collect new centroid locations and iterated thousands of times until a significant fraction of fluorescent label molecules have been sampled. (See the movie for an illustration of this principle.)

Figure 1A: A high-resolution optical microscope image of a fluorescently labeled Golgi in a cell, demonstrating the limits of optical resolution. 1B: The same area imaged and processed by the PALM approach, resulting in significantly higher resolution.

If we draw only the centers (and not the whole fuzzy ball) of all these fluorescent molecules that have each been imaged individually, we can generate a high-resolution image. The images in Figure 1 show a fluorescent protein–labeled Golgi apparatus in a cell seen by regular microscopy and by PALM.

With colleagues at the National Institutes of Health and Florida State University, we are broadening the utility of this technique and demonstrating ever more compelling applications to biological problems. Developing and streamlining sample preparation techniques, that preserve cell structure and optimize photophysical properties of fluorescent labels, is key for expanding the application. Furthermore, instead of labeling only one protein, two or even more proteins can be labeled, each with a distinguishing trait such as color. This could paint a picture of how one protein works in relation to another.

Interferometry is a technique that can measure positions to nanometer accuracy. To gain access to the third axial dimension, we have recently combined interferometry with PALM in a method we call iPALM that can resolve individual fluorescent protein locations in three dimensions. Requirements of self-calibration and tolerance to fluctuating brightness of the labels can be met with simultaneous multiphase interferometry and inspired the invention of a custom three-way beam splitter. Figure 2 illustrates how light from a single-molecule source interferes into three different beams, depending on the axial location of the molecule. The nanometer accuracy can decipher the three-dimensional structure of protein complexes such as focal adhesion, an assembly of more than 100 proteins that the cell uses to attach its cytoskeleton to an external environment and is used in motion or to respond to forces. (For more details, see Super-Resolution Microscopy Takes on a Third Dimension.) The first comprehensive application of iPALM was to tease apart the molecular architecture of a cellular structure called focal adhesions. These ~ 100 nm thick plaques consist of over 100 different proteins that connect an object outside the cell, in our case the glass slide surface, through the membrane to the actin fibers of the cytoskeleton. With data from the iPALM we were able to see the layering of the different proteins and in certain cases even determine the orientation of longer proteins.

PALM and iPALM are only initial examples of what I hope will be a series of broadly useful innovations in microscopy. We rely on outside researchers to define and explore promising applications of biology to the iPALM instrument and welcome potential collaborations.

Biological systems are intrinsically three dimensional. Capturing this third dimension with both good fidelity and high throughput is particularly useful for neural circuit reconstruction. Electron microscopes are the time-tested way to resolve the thin membranes, synapse, and fine processes that define a circuit. There are different electron microscope technologies, each with advantages and trade-offs in throughput, lateral resolution, vertical resolution, defects of missing volumes, and ease of automation. Furthermore, since the true limitations are in image processing, these data attributes critically influence ease of registration, segmentation, tracing, annotation, etc. Obtaining the usable data requires a tight interaction between sample preparation, image processing, and data acquisition for best and timely reconstruction. Sample preparation, algorithms, and acquisition development go hand in hand.

We are optimizing and evaluating two approaches for this specific application: tilt STEM (scanning transmission electron microscopy) and FIB-SEM (focused ion beam with scanning electron microscopy). Since each transmitted primary electron can impart a greater signal-to-noise ratio than backscattered electrons, transmission microscopy offers a greater throughput for a given number of primary electrons. Furthermore, STEM allows one to scan large areas with fine pixilation (minimizing the need to splice multiple images), can dynamically focus on tilted surfaces (which is required on large areas), and is a convenient platform for automation. The sections used in transmission are cut 40–50 nm, which defines the z resolution for normal imaging with serial sections. This is inadequate; however, tilted imaging, at a few angles, can give valuable depth resolution within one section. How this projects out to reconstruction quality again rests on the interplay between data acquisition parameters and algorithms for processing data, and we are exploring this with Mitya Chklovskii.

Focused ion beams were developed more than 20 years ago for fine cross-sectioning in materials science and for semiconductor chip analysis. A fine atomic beam with <10-nm diameter mills out trenches or polishes walls. These newly exposed surfaces, such as a cross section of a transistor, can then be imaged with a scanning electron microscope (SEM) that detects the scattered electrons. This is now being applied to biological samples, particularly neural circuits. A sequence of fine polishing steps of 10 nm or less, each followed by imaging of each new surface, can give a stack of 3D data with isotropic resolution. Such continuous milling/imaging also gives excellent registration and avoids many of the defects, such as tears and folds associated with cutting the thin sections required for transmission microscopy. However, this comes at the cost of slower imaging speeds. Again, a tight interaction from sample preparation to image processing is evolving this approach and defining how it will best serve open biological questions.

Many biomolecules in cells can be visualized with high sensitivity and specificity by fluorescence microscopy. However, the resolution of conventional light microscopy is limited by diffraction to ~200-250nm laterally and >500nm axially. Here, we describe superresolution methods based on single-molecule localization analysis of photoswitchable fluorophores (PALM: photoactivated localization microscopy) as well as our recent three-dimensional (3D) method (iPALM: interferometric PALM) that allows imaging with a resolution better than 20nm in all three dimensions. Considerations for their implementations, applications to multicolor imaging, and a recent development that extend the imaging depth of iPALM to ~750nm are discussed. As the spatial resolution of superresolution fluorescence microscopy converges with that of electron microscopy (EM), direct imaging of the same specimen using both approaches becomes feasible. This could be particularly useful for cross validation of experiments, and thus, we also describe recent methods that were developed for correlative superresolution fluorescence and EM.

The human immunodeficiency virus (HIV) hijacks the endosomal sorting complexes required for transport (ESCRT) to mediate virus release from infected cells. The nanoscale organization of ESCRT machinery necessary for mediating viral abscission is unclear. Here, we applied three-dimensional superresolution microscopy and correlative electron microscopy to delineate the organization of ESCRT components at HIV assembly sites. We observed ESCRT subunits localized within the head of budding virions and released particles, with head-localized levels of CHMP2A decreasing relative to Tsg101 and CHMP4B upon virus abscission. Thus, the driving force for HIV release may derive from initial scaffolding of ESCRT subunits within the viral bud interior followed by plasma membrane association and selective remodeling of ESCRT subunits.

We combine super-resolution localization fluorescence microscopy with transmission electron microscopy of metal replicas to locate proteins on the landscape of the cellular plasma membrane at the nanoscale. We validate robust correlation on the scale of 20 nm by imaging endogenous clathrin (in two and three dimensions) and apply the method to find the previously unknown three-dimensional position of the endocytic protein epsin on clathrin-coated structures at the plasma membrane.

The ability to localize proteins precisely within subcellular space is crucial to understanding the functioning of biological systems. Recently, we described a protocol that correlates a precise map of fluorescent fusion proteins localized using three-dimensional super-resolution optical microscopy with the fine ultrastructural context of three-dimensional electron micrographs. While it achieved the difficult simultaneous objectives of high photoactivated fluorophore preservation and ultrastructure preservation, it required a super-resolution optical and specialized electron microscope that is not available to many researchers. We present here a faster and more practical protocol with the advantage of a simpler two-dimensional optical (Photoactivated Localization Microscopy (PALM)) and scanning electron microscope (SEM) system that retains the often mutually exclusive attributes of fluorophore preservation and ultrastructure preservation. As before, cryosections were prepared using the Tokuyasu protocol, but the staining protocol was modified to be amenable for use in a standard SEM without the need for focused ion beam ablation. We show the versatility of this technique by labeling different cellular compartments and structures including mitochondrial nucleoids, peroxisomes, and the nuclear lamina. We also demonstrate simultaneous two-color PALM imaging with correlated electron micrographs. Lastly, this technique can be used with small-molecule dyes as demonstrated with actin labeling using phalloidin conjugated to a caged dye. By retaining the dense protein labeling expected for super-resolution microscopy combined with ultrastructural preservation, simplifying the tools required for correlative microscopy, and expanding the number of useful labels we expect this method to be accessible and valuable to a wide variety of researchers.

A central problem in neuroscience is reconstructing neuronal circuits on the synapse level. Due to a wide range of scales in brain architecture such reconstruction requires imaging that is both high-resolution and high-throughput. Existing electron microscopy (EM) techniques possess required resolution in the lateral plane and either high-throughput or high depth resolution but not both. Here, we exploit recent advances in unsupervised learning and signal processing to obtain high depth-resolution EM images computationally without sacrificing throughput. First, we show that the brain tissue can be represented as a sparse linear combination of localized basis functions that are learned using high-resolution datasets. We then develop compressive sensing-inspired techniques that can reconstruct the brain tissue from very few (typically 5) tomographic views of each section. This enables tracing of neuronal processes and, hence, high throughput reconstruction of neural circuits on the level of individual synapses.

The molecular mechanism responsible for capturing, sorting and retrieving vesicle membrane proteins following triggered exocytosis is not understood. Here we image the post-fusion release and then capture of a vesicle membrane protein, the vesicular acetylcholine transporter, from single vesicles in living neuroendocrine cells. We combine these measurements with super-resolution interferometric photo-activation localization microscopy and electron microscopy, and modelling to map the nanometer-scale topography and architecture of the structures responsible for the transporter's capture following exocytosis. We show that after exocytosis, the transporter rapidly diffuses into the plasma membrane, but most travels only a short distance before it is locally captured over a dense network of membrane-resident clathrin-coated structures. We propose that the extreme density of these structures acts as a short-range diffusion trap. They quickly sequester diffusing vesicle material and limit its spread across the membrane. This system could provide a means for clathrin-mediated endocytosis to quickly recycle vesicle proteins in highly excitable cells.

Microscopic images of specific proteins in their cellular context yield important insights into biological processes and cellular architecture. The advent of superresolution optical microscopy techniques provides the possibility to augment EM with nanometer-resolution fluorescence microscopy to access the precise location of proteins in the context of cellular ultrastructure. Unfortunately, efforts to combine superresolution fluorescence and EM have been stymied by the divergent and incompatible sample preparation protocols of the two methods. Here, we describe a protocol that preserves both the delicate photoactivatable fluorescent protein labels essential for superresolution microscopy and the fine ultrastructural context of EM. This preparation enables direct 3D imaging in 500- to 750-nm sections with interferometric photoactivatable localization microscopy followed by scanning EM images generated by focused ion beam ablation. We use this process to "colorize" detailed EM images of the mitochondrion with the position of labeled proteins. The approach presented here has provided a new level of definition of the in vivo nature of organization of mitochondrial nucleoids, and we expect this straightforward method to be applicable to many other biological questions that can be answered by direct imaging.

A fundamental objective in molecular biology is to understand how DNA is organized in concert with various proteins, RNA, and biological membranes. Mitochondria maintain and express their own DNA (mtDNA), which is arranged within structures called nucleoids. Their functions, dimensions, composition, and precise locations relative to other mitochondrial structures are poorly defined. Superresolution fluorescence microscopy techniques that exceed the previous limits of imaging within the small and highly compartmentalized mitochondria have been recently developed. We have improved and employed both two- and three-dimensional applications of photoactivated localization microscopy (PALM and iPALM, respectively) to visualize the core dimensions and relative locations of mitochondrial nucleoids at an unprecedented resolution. PALM reveals that nucleoids differ greatly in size and shape. Three-dimensional volumetric analysis indicates that, on average, the mtDNA within ellipsoidal nucleoids is extraordinarily condensed. Two-color PALM shows that the freely diffusible mitochondrial matrix protein is largely excluded from the nucleoid. In contrast, nucleoids are closely associated with the inner membrane and often appear to be wrapped around cristae or crista-like inner membrane invaginations. Determinations revealing high packing density, separation from the matrix, and tight association with the inner membrane underscore the role of mechanisms that regulate access to mtDNA and that remain largely unknown.

We combined photoactivated localization microscopy (PALM) with live-cell single-particle tracking to create a new method termed sptPALM. We created spatially resolved maps of single-molecule motions by imaging the membrane proteins Gag and VSVG, and obtained several orders of magnitude more trajectories per cell than traditional single-particle tracking enables. By probing distinct subsets of molecules, sptPALM can provide insight into the origins of spatial and temporal heterogeneities in membranes.

Commentary: As a stepping stone to true live cell PALM (see above), our collaborator Jennifer Lippincott-Schwartz suggested using the sparse photoactivation principle of PALM to track the nanoscale motion of thousands of individual molecules within a single living cell. Termed single particle tracking PALM (sptPALM), Jennifer’s postdocs Suliana Manley and Jen Gillette used the method in our PALM rig to create spatially resolved maps of diffusion rates in the plasma membrane of live cells. sptPALM is a powerful tool to study the active cytoskeletal or passive diffusional transport of individual molecules with far more measurements per cell than is possible without sparse photoactivation.

We introduce a method for optically imaging intracellular proteins at nanometer spatial resolution. Numerous sparse subsets of photoactivatable fluorescent protein molecules were activated, localized (to approximately 2 to 25 nanometers), and then bleached. The aggregate position information from all subsets was then assembled into a superresolution image. We used this method--termed photoactivated localization microscopy--to image specific target proteins in thin sections of lysosomes and mitochondria; in fixed whole cells, we imaged vinculin at focal adhesions, actin within a lamellipodium, and the distribution of the retroviral protein Gag at the plasma membrane.

Commentary: The original PALM paper by myself and my friend and co-inventor Harald Hess, spanning the before- and after-HHMI eras. Submitted and publicly presented months before other publications in the same year, the lessons of the paper remain widely misunderstood: 1) localization precision is not resolution; 2) the ability to resolve a few molecules by the Rayleigh criterion in a diffraction limited region (DLR) does not imply the ability to resolve structures of arbitrary complexity at the same scale; 3) true resolution well beyond the Abbe limit requires the ability to isolate and localize hundreds or thousands of molecules in one DLR; and 4) certain photoactivatable fluorescent proteins (PA-FPs) and caged dyes can be isolated and precisely localized at such densities; yielding true resolution down to ~20 nm. The molecular densities we demonstrate (105 molecules/m2) are more than two orders of magnitude greater than in later papers that year (implying ten-fold better true resolution) – indeed, these papers demonstrate densities only comparable to earlier spectral or photobleaching based isolation methods. We validate our claims by correlative electron microscopy, and demonstrate the outstanding advantages of PA-FPs for superresolution microscopy: minimally perturbative sample preparation; high labeling densities; close binding to molecular targets; and zero non-specific background.

Luminescent centers with sharp (<0.07 millielectron volt), spectrally distinct emission lines were imaged in a GaAs/AIGaAs quantum well by means of low-temperature near-field scanning optical microscopy. Temperature, magnetic field, and linewidth measurements establish that these centers arise from excitons laterally localized at interface fluctuations. For sufficiently narrow wells, virtually all emission originates from such centers. Near-field microscopy/spectroscopy provides a means to access energies and homogeneous line widths for the individual eigenstates of these centers, and thus opens a rich area of physics involving quantum resolved systems.